Compendium of Pea Diseases and Pests, Third Edition

seed to seedling: Effects of inoculum dose, inoculation method, temperature and soil moisture. J. Appl. Bacteriol. 81:65-72.

Skoric, V. 1927. Bacterial blight of pea: Overwintering, dissemination, and pathological history. Phytopathology 17:611- 627.

Susuri, L., Hagedorn, D. J., and Rand, R. E. 1982. Alternaria blight of pea. Plant Dis. 66:328-330.

Taylor, J. D., and Dye, D. W. 1972. A survey of the organisms associated with bacterial blight of peas. N. Z. J. Agric. Res. 15:432- 440.

Taylor, J. D., and Dye, D. W. 1976. Evaluation of streptomycin seed treatments for the control of bacterial blight of peas (Pseudomonas pisi Sackett 1916). N. Z. J. Agric. Res. 19:91-95.

Teixeira, C. S. P. 2005. Are Stemphylium spp. seed borne pathogens of pea (Pisum sativum L.)? M.S. thesis. Lincoln University, Christchurch, New Zealand.

Tenuta, M., Madani, M., Briar, S., Molina, O. I., Gulden, R., and Subbotin, S. A. 2015. Occurrence of Ditylenchus weischeri and not D. dipsaci in field pea harvest samples and Cirsium arvense in the Canadian prairies. J. Nematol. 46:376-384.

Timmerman-Vaughn, G., Larsen, R., Murray, S., McPhee, K., and Coyne, C. 2009. Analysis of the accumulation of Pea enation mosaic virus genomes in seed tissues and lack of evidence for seed transmission in pea (Pisum sativum). Phytopathology 99:1281-1288.

Willman, K., Stepien, L., Fabianska, I., and Kachlicki, P. 2014. Plantpathogenic fungi in seeds of different pea cultivars in Poland. Arch.

Ind. Hyg. Toxicol. 65:329-338.

(Prepared by B. Agindotan and M. Burrows)

Bacterial Diseases Bacterial Blight In the past, the disease name “bacterial blight” of pea referred to infection caused by Pseudomonas syringae pv. pisi. In more recent times, P. syringae pv. syringae (syn. P. syringae) has also been increasingly implicated with the disease in Spain and Australia. P. viridiflava, P. fluorescens, and soft-rotting pseudomonads have also been isolated from pea with bacterial blight symptoms. Similarly, symptoms of brown spot of pea can also be caused by P. syringae pv. syringae and P. syringae pv. pisi. Hence, the disease name bacterial blight can be applied to a complex of P. syringae pv. syringae and P. syringae pv. pisi that causes disease symptoms on pea, including those symptoms previously referred to separately as bacterial blight and brown spot of pea.

Since the first description of bacterial blight in Colorado in 1915 by W. G. Sackett, this seed- and stubble-borne disease has been found in other areas of the United States and Canada, Bermuda, South America (Argentina and Colombia), Africa (Kenya, Lesotho, Morocco, South Africa, and Tanzania), Asia (India, Indonesia, Japan, Lebanon, Nepal, and Pakistan), Australia and New Zealand, and Europe (Bulgaria, France, Germany, Greece, Hungary, Italy, the Netherlands, Portugal, Romania, Spain, and the United Kingdom). P. syringae pv. syringae can be found in most areas where pea is grown and was described on pea in the United States in Wisconsin in 1966 and Idaho in 1970 and in New Zealand in 1972. It can be found more commonly in the Northern Hemisphere on pea planted in the autumn or winter.

Bacterial blight can be sporadic and quite severe (Fig. 1), particularly during seasons with frost damage, high humid-

Fig. 1. Pea field severely affected by bacterial blight after a hailstorm. (Courtesy R. M. Harveson—© APS)

ity, and wet conditions or when overhead sprinkler irrigation is used. Yield losses are strongly correlated with disease severity (approximately 0.5 t/ha for each 10% increase in affected canopy area).

Symptoms Bacterial blight symptoms can be found on all aboveground plant tissues. Initial symptoms appear as small, shiny, watersoaked lesions on leaves. Lesions eventually coalesce, turning brown and necrotic, and may take on an angular appearance (Fig. 2). Leaves “burned” by infection can soon dry and drop off, and no further disease progression occurs unless a humid environment is provided.

Stem infections appear as irregular flecks around the nodes, stipules, and leaflets. These lesions enlarge and coalesce, becoming sunken and elongate (Fig. 3). The lesions often girdle the stem, causing death of the plant above the infected area.

Stem lesions can severely distort the stems, petioles, and growing points. Infection of a stipule usually starts at the point of attachment to the stem. The interveinal tissue first becomes water soaked and then yellow and later brown and papery. The spread of the pathogen in stipules is delimited by the veins, which results in the characteristic fanlike lesions of a bacterial blight infection (Fig. 4). Prevailing weather conditions dictate the severity of symptom development. Dry conditions with occasional frosts increase disease progression and the level of plant injury. Rainy, wet conditions result in less severe crop injury but can increase the rate of disease spread.

Infection of floral parts may also occur and generally leads to stem and pod infections. Under extremely moist conditions, droplets of bacterial ooze may be observed from infected tissue

Fig. 2. Pea leaf with bacterial blight lesions coalescing and becoming angular. (Courtesy R. M. Harveson—© APS)

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or dried on the surface tissue (Figs. 5 and 6). Flower buds may appear shriveled and decayed when sepals are infected early in the blossom stage. Later infections may lead to pod infection, resulting in water-soaked, sunken lesions (Fig. 7). Seed within these pods may be covered with bacterial slime that dries to a thin, white film. Bacteria may occasionally penetrate the pod and damage the seed. The disease is most evident in commercial crops when patches of dead plants, 1–2 m in diameter,

develop. Symptoms can be easily confused with those of Ascochyta blight (Fig. 8), but Ascochyta lesions are not limited by veins, may have pycnidia associated with the lesions, and are not associated with patterns caused by hail such as damage on the side of the plant exposed to the prevailing wind. Coinfections of the same plant commonly occur.

Causal Organisms P. syringae pv. pisi and P. syringae pv. syringae are both aerobic, gram-negative, non-spore-forming, motile rods (average size 0.7 × 2–3 µm) with one to several polar flagella.

On King’s B medium, P. syringae pv. syringae isolates produce pigments more rapidly and in larger quantities than do

Fig. 3. Advanced bacterial blight lesions on a pea stem are elongate, dried, and sunken. (Cour tesy R. M. Harveson— © APS)

Fig. 6. Dried bacterial growth on a pea leaf. (Cour tesy M. Burrows— © APS)

Fig. 4. Dried, fanlike bacterial blight lesions between the veins of a pea leaf. (Cour tesy R. M. Harveson— © APS) Fig. 7. Dried, sunken bacterial blight lesions on pea pods. (Courtesy R. M. Harveson— © APS)

Fig. 5. Bacterial growth oozing from lesions under conditions of high humidity. (Courtesy M. Burrows—© APS)

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Fig. 8. Pea leaf with a bacterial blight lesion (right) compared with the more circular lesion characteristic of Ascochyta blight (left).

(Courtesy M. Burrows—© APS)

P. syringae pv. pisi isolates. P. syringae pv. syringae pigment is blue green compared with the green hue characteristic of those races of P. syringae pv. pisi that fluoresce (not all isolates of pathovar pisi fluoresce). Acrylamide electrophoretic protein patterns of the two differ in at least two bands. Differences in peroxidase isozymes have also been demonstrated. P. syringae pv. syringae produces a toxin (syringomycin) in culture that inhibits growth of Geotrichum candidum; P. syringae pv. pisi does not. The two pathovars can be differentiated by phage and serology tests as well as by pathogenicity tests conducted on pea and bean. P. syringae pv. pisi does not infect bean.

These bacteria belong in group Ia of the fluorescent plantpathogenic pseudomonads. Members of this group are all characterized as levan production positive, oxidase negative, potato rot negative, and arginine dihydrolase negative and cause a hypersensitive reaction in tobacco (i.e., LOPAT: +–––+). P. syringae pv. pisi can be readily distinguished from P. syringae pv. syringae by standard physiological tests. All strains of P. syringae pv. pisi are negative for gelatin liquefaction and can utilize homoserine. In contrast, P. syringae pv. syringae is positive for gelatin liquefaction and cannot utilize homoserine. P. syringae pv. pisi also can be distinguished from other fluorescent pseudomonads on the basis of the pathogenic reaction on pea.

Inoculation on pods or leaves initially results in water-soaked spots, whereas inoculation with P. syringae pv. syringae produces necrotic spots. Other fluorescent bacteria produce no symptoms.

In early studies, bacteriophage typing and serological tests were used to identify these pathogens, but current polymerase chain reaction (PCR) tests allow differentiation between P. syringae pv. syringae and P. syringae pv. pisi.

At present, eight races of P. syringae pv. pisi have been described on the basis of pathogenicity to differential cultivars.

A gene-for-gene relationship exists between the pathogen avirulence (avr) gene and the host resistance gene. Results from several studies in which the distribution of races in over 300 isolates was investigated identified only one isolate of race 7, eight of race 1, and a low frequency of race 5. Each of these races contains multiple avirulence genes. Race 6, which lacks all avirulence genes, was reported in all studies conducted, with more recent studies suggesting that the frequency of this race may be increasing. Several studies have shown that the distribution of races, other than race 6, is associated with the resistance genes present in the predominant cultivars grown in a region. For example, the dominance of race 2 (82%) in the United Kingdom was attributed to the low frequency of resistance gene 2 in the dominant pea cultivars, while race 3 (64%) was most common in Australia, which was attributed to the dominance of the race 3-susceptible cultivars being grown.

Disease Cycle and Epidemiology Both P. syringae pv. pisi and P. syringae pv. syringae can be transmitted by seed. P. syringae pv. pisi is carried both externally and internally and is known to persist for 3 years. In contrast, P. syringae pv. syringae is carried primarily externally and has a shorter survival time on seed.

When pods are severely infected by P. syringae pv. pisi, the bacterium can infect the funiculus and enter mature seed via the micropyle. Populations then colonize the intercellular and intracellular spaces of the seed coat. Because the bacterium does not penetrate the embryo or cotyledons, infected seeds remain viable. Seed infestation levels can vary depending on seed maturity, weather conditions, and the severity of pod infection.

During germination, the plumule becomes infected when it contacts the infected seed coat. Transmission level from infected seeds to plants is lower in dry than in moist soils. Under high moisture conditions, the entire crop may be lost. Under less optimal conditions, growing tips may be killed by blight and new stems may arise from lower nodes. In the absence of favorable environmental conditions, infected seedlings may go

unnoticed. Lower leaves that are infected are covered by the canopy. These infected leaves can dry and become a fine chaff that later becomes a source of contamination when seeds are processed.

P. syringae pv. pisi is a strong colonizer of plant surfaces.

This epiphytic nature contributes to the accumulation and maintenance of high bacterial populations in the absence of symptoms and can facilitate spread within a crop when favorable conditions occur, in particular wind and rain or frost injury. Populations ranging from 10 6 to 108 colony-forming units per gram of fresh weight can be found on both resistant and susceptible cultivars of pea. Weeds present in infected fields harbor slightly lower populations of the pathogen.

Residues from an infected pea crop can be an important source of inoculum for both P. syringae pv. pisi and P. syringae pv. syringae, and crop rotation is considered an important management strategy. Studies in Australia showed that P. syringae pv. pisi could survive on residues in the field for up to 104 weeks and P. syringae pv. syringae for up to 34 weeks and that these survival times could be greatly reduced with burial of crop residues. Since survival time of the bacteria in soil is very short, it is not considered an important source of infection for a new crop.

Mechanical injury can predispose a pea crop to bacterial blight infection through the creation of wounds in plants.

Wounds may be induced by hail, sandstorms, wind, rain, or cultural practices. Frost events are also closely associated with bacterial blight outbreaks because of the ice-nucleating properties of P. syringae. It is thought that P. syringae has evolved the capacity to predispose plant tissue to frost damage and subsequent bacterial penetration and disease development, thus conferring a competitive survival advantage.

The bacterium’s epiphytic survival ability is evidenced by the large number of host plants that are colonized. Alternative hosts include sweet pea, everlasting pea, cowpea, hyacinth bean, purple vetch, hairy vetch, red clover, and soybean. In the case of red clover and soybean, only P. syringae pv. pisi race 2 is pathogenic. Whether separate races have been used to test pathogenicity to the other species noted is not known. Other plants that host epiphytic populations of P. syringae pv. pisi are felon herb, mugwort, wormweed, slender-flower thistle, carrot, bacon weed, fat-hen, goosefoot, lamb’s-quarter, meal-weed, pigweed, white turtlehead, Bermudagrass, devil’s grass, dog’s tooth grass, scotch grass, wire grass, Hopi sunflower, mirasol, sunflower, wild sunflower, bird’s-foot trefoil, black medic, hop clover, nonesuch clover, yellow trefoil, melist, yellow clover, melilot yellow sweet clover, primrose, groundsel, California rape, charlock, Irish potato, potato, white potato, black-wort, boneset, bruisewort, comfrey, ivy-leaved speedwell, dent corn, field corn, flint corn, maize, pod corn, popcorn, sweet corn, and volunteer corn. P. syringae pv. syringae has a wide range of other host plants, including citrus, chrysanthemum, bean, hibiscus, walnut, apple, Pennisetum spp., pepper, plum, pear, rose, grain, lilac, clover, vetch, and corn.

Management A range of methods can contribute to the management of bacterial blight of pea, including planting disease-free seed, avoiding fields where infested crop residues are present, timing of sowing and avoidance of frost-prone fields, disinfection of equipment, and use of resistant cultivars.

The simplest method is the use of resistant cultivars. As mentioned previously, a gene-for-gene relationship exists between the host and P. syringae pv. pisi. Single or multiple avirulence genes have been found in all races except for race 6, to which all current Pisum sativum accessions are susceptible. Resistance to P. syringae pv. syringae in field pea shows an almost continuous disease response, suggesting polygenic resistance to this pathovar. Resistance to this pathovar has been reported in Australia and Spain. Numerous commercial field pea cultivars 11

are available worldwide with resistance to P. syringae pv. syringae.

Planting clean seed is also an important means of managing bacterial blight. To reduce seed contamination, it is helpful to produce seed in arid areas, avoid sprinkler irrigation, and inspect fields for symptoms. Farm and seed-processing equipment should be cleaned between seed lots, and early-season planting should be avoided in order to minimize injury from unfavorable weather conditions. P. syringae pv. pisi is a strong epiphyte; therefore, field inspections may not identify infected seed lots. Seed health testing for the pathogen is recommended to ensure quality. At present, there is no prescribed quantity of seed that should be assayed for the pathogen. However, assays for a similar bacterium, P. syringae pv. phaseolicola, require testing 5,000 seeds of each lot. If the pathogen is detected on seed, use of the seed should be deferred for 1 year.

Infested crop residues have been shown to be important inoculum sources for both P. syringae pv. pisi and P. syringae pv. syringae. Studies show that survival is reduced by burial of crop residues and recommend that there should be at least a 2-year break following an infected crop. Experimentally, treatment of seed with chemicals (e.g., sodium hypochlorite or streptomycin sulfate) was found to be only partially effective.

Good results were obtained with dry heat, hot-water soaks, and a combination of these treatments, but they are not considered to be commercially viable.

Disinfestation of harvesting equipment between field sites is recommended. During the season, walking through fields when the vines are wet should be avoided. A good practice is to avoid traveling from infested fields to clean fields.

Symptoms On pea seed, the name of the disease is descriptive, although the degree of pigmentation is variable (Fig. 9). It is this variability, plus its permanence, that readily distinguishes diseased seed from seed coated with a pink-colored pesticide that is easily removed by scrubbing in water. On pea pods, discoloration is less intense, tending to a brownish pink rather than bright pink discoloration (Fig. 10). The bacterium responsible for pink seed can also induce a crown and shoot rot of pea, with pinkish to dark brownish black discoloration of tissues.

Causal Organism Erwinia rhapontici is a gram-negative bacterium producing pinkish colonies on artificial media such as sucrose peptone agar but is less reliably pinkish on potato dextrose agar. It is a member of the carotovora (soft rot) group in Erwinia but degrades various sugars rather than pectate.

Management E. rhapontici has demonstrated potential for seed transmission. Strains originating from pulses can infect other pulses as well as cereals, and strains from cereals can infect pulses.

Other than exclusion of symptomatic seed lots from plantings, the minor importance of pink seed has thus far precluded development of management strategies specifically focused on this pathogen. However, grain inspection authorities have noted that symptoms of pink seed are similar to those of pink-colored pesticide seed treatments, so correct diagnosis can be essential to avoid mistakenly labeling seed lots as pesticide contaminated.

Selected References Bevan, J. R., Taylor, J. D., and Crute, I. R. 1995. Genetics of specific resistance in pea (Pisum sativum) cultivars to seven races of Pseudomonas syringae pv. pisi. Plant Pathol. 44:98-108.

Grondeau, C., Mabiala, A., Ait-Oumeziane, R., and Samson, R. 1996.

Epiphytic life is the main characteristic of the life of Pseudomonas syringae pv. pisi, pea bacterial blight agent. Eur. J. Plant Pathol.

102:353-363.

Hollaway, G. J., Bretag, T. W., and Price, T. V. 2007. The epidemiology and management of bacterial blight (Pseudomonas syringae pv. pisi) of field pea (Pisum sativum) in Australia: A review. Aust.

J. Agric. Res. 58:1086-1099.

Richardson, H. J., and Hollaway, G. J. 2011. Bacterial blight caused by Pseudomonas syringae pv. syringae shown to be an important disease of field pea in south eastern Australia. Australas. Plant Pathol.

40:260-268.

Sackett, W. G. 1916. A bacterial stem blight of field and garden peas.

Colo. Agric. Exp. Stn. Bull. 218:3- 43.

Taylor, J. D., and Dye, D. W. 1972. A survey of the organisms associated with bacterial blight of peas. N. Z. J. Agric. Res. 15:432- 440.

Taylor, J. D., Bevan, J. R., Crute, I. R., and Reader, S. L. 1989. Genetic relationship between races of Pseudomonas syringae pv. pisi and cultivars of Pisum sativum. Plant Pathol. 38:364-375.

Fig. 9. Pink seed. Symptomatic field pea seeds with variable distribution of pink pigment on seed coats. (Cour tesy S. Lupien)

(Prepared by K. D. Lindbeck, G. J. Hollaway, F. Mathew, and R. M. Harveson)

Pink Seed Pink seed of field pea (Pisum sativum) was first reported in North America from Canada and subsequently from the United States. Pink seed is also known to occur on chickpea (Cicer arietinum), lentil (Lens culinaris), and bean (Phaseolus vulgaris).

It has long been known in grains of cereals, especially durum and bread wheat (Triticum spp.). The bacterium that causes pink seed is also of minor or sporadic importance in some other crops such as rhubarb (Rheum rhaponticum), garlic and onion (Allium spp.), and hyacinth (Hyacinthus orientalis).

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Fig. 10. Pink seed on field pea pods. Left, three asymptomatic pods, pale green to green; right, five symptomatic pods with brownish pink patches. (Cour tesy S. Lupien)

Selected References Adesemoye, A. O., Wei, H. H., and Harveson, R. M. 2016. Identification of Erwinia rhapontici as the causal agent of crown and shoot rot and pink seed of pea in Nebraska. Plant Health Prog. 17:155-157.

Dugan, F. M., Lupien, S. L., and Schroeder, B. K. 2003. Pink seed of pea. Wash. State Univ., Ext. Serv., Ext. Bull. 1967.

Fusikovsky, L. A. 2010. Pink seed. Page 12 in: Compendium of Wheat Diseases and Pests. 3rd ed. W. W. Bockus, R. L. Bowden, R. M. Hunger, W. L. Morrill, T. D. Murray, and R. W. Smiley, eds. American Phytopathological Society, St. Paul, MN. Hsieh, T. F., Huang, H. C., and Erickson, R. S. 2010. Spread of seedborne Erwinia rhapontici in bean, pea and wheat. Eur. J. Plant Pathol. 127:579-584.

Huang, H. C., Erickson, R. S., and Hsieh, T. F. 2007. Lack of host specificity of strains of Erwinia rhapontici, causal agent of pink seed of pulse and cereal crops. Bot. Stud. 48:181-186.

Huang, H. C., Erickson, R. S., Yanke, L. J., Hsieh, T. F., and Morrall, R. A. A. 2003. First report of pink seed of lentil and chickpea caused by Erwinia rhapontici in Canada. Plant Dis. 87:1398.

Huang, H. C., Phillippe, L. M., and Phillippe, R. C. 1990. Pink seed of pea: A new disease caused by Erwinia rhapontici. Can. J. Plant Pathol. 12:445.

Schroeder, B. K. 2011. Diseases caused by bacteria and a phytoplasma.

Pages 91-94 in: Compendium of Chickpea and Lentil Diseases and Pests. W. Chen, H. C. Sharma, and F. J. Muehlbauer, eds. American Phytopathological Society, St. Paul, MN. Schroeder, B. K., Lupien, S. L., and Dugan, F. M. 2002. First report of pink seed of pea caused by Erwinia rhapontici in the United States.

Plant Dis. 86:188.

(Prepared by F. M. Dugan, R. M. Harveson, and H. C. Huang)

Fungal and Oomycete Diseases Root Diseases Aphanomyces Root Rot Aphanomyces root rot, also called common root rot, is a widespread and destructive disease of pea that develops primarily in wet soil. Aphanomyces root rot is found in many peagrowing areas of North America, northern Europe, Australia, New Zealand, and Japan. In the United States, this disease is very damaging to pea in the Great Lakes, Northeastern, and Pacific Northwest regions. It is an emerging concern in the peaproducing areas of North Dakota and Montana and in the Canadian prairies, where it was first reported in 2013 and 2014. In Europe, the disease has caused significant losses on pea crops, particularly in France and Scandinavia, since the mid 1990s.

Disease severity and yield losses are most severe when soil is wet and warm. Entire fields can be destroyed under favorable conditions. In many pea-production areas in the United States, Canada, and Europe, Aphanomyces root rot is considered the most important disease of pea.

Fig. 11. Aphanomyces root rot. (Courtesy S. Chatterton—© Her Majesty the Queen in Right of Canada, as represented by the Minister of Agriculture and Agri-Food Canada)

Symptoms Symptoms of Aphanomyces root rot appear on the roots within 2 weeks after pathogen infection. Infected areas are initially honey brown, and cortical tissue becomes soft and darker brown as the disease progresses (Fig. 11). When infected plants are pulled from the soil, the root cortex often sloughs off and remains behind in the soil, leaving only the central strand of vascular tissue attached to the rest of the plant (Fig. 12). Microscopic observation usually reveals oospores in infected cortical tissue (Fig. 13). Soon after symptoms develop on roots, the infection moves up into the epicotyl, which initially becomes honey colored and soft. The epicotyl darkens as other organisms invade the infected tissue and often becomes shrunken and completely rotted in advanced stages of disease. Plants may become severely wilted, chlorotic, and stunted if infection occurs early in the season. These symptoms and plant stunting are

Fig. 12. Advanced symptoms of Aphanomyces root rot. The root cortex sloughs off, leaving the inner bundle of vascular tissue.

(Courtesy S. Chatterton—© Her Majesty the Queen in Right of Canada, as represented by the Minister of Agriculture and AgriFood Canada)

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further exacerbated by a reduction in the number of rhizobial nodules. Leaves of an infected plant yellow progressively from the bottom of the shoot upward (Fig. 14). Pod production is reduced, pods may contain reduced numbers of seeds, or plants may die before pod development. The symptoms and yield loss depend largely on the environment, which influences time and severity of infection and the degree of water stress in diseased plants.

Causal Organism Aphanomyces root rot is caused by a funguslike organism, Aphanomyces euteiches, an oomycete now classified in the kingdom Stramenopila (not true fungi). Two kinds of reproductive spores are produced, oospores and zoospores (Fig. 15).

The thick-walled oospores (18–25 µm in diameter) develop in infected root cortical tissue and are released into soil from decomposing roots (Fig. 13). The oospores are believed to germinate in response to chemical compounds released by roots and then form either hyphae or zoosporangia, depending in part on the nutrient status of the soil. The zoosporangia, which are morphologically undifferentiated from the hyphae, produce asexual swimming spores called zoospores (8–12 µm in diameter). Zoosporangia can arise from hyphae or germinating oospores. The primary asexual zoospores emerge and aggregate into grapelike clusters at the apex of zoosporangia (Fig. 16).

Motile zoospores (Fig. 15) emerge from these primary spores and are attracted to roots by unidentified chemical signals released from roots. After swimming for a brief time with the aid of two flagella, the zoospores encyst and germinate on the surfaces of roots. Hyphae grow from the encysted zoospores, penetrate the roots, grow through the roots, and release enzymes that break down host tissue.

A. euteiches can be induced to produce zoospores and oospores in pure culture. Good mycelial growth and oospore production can be achieved on cornmeal agar, where the pathogen produces sparse growth within and on the surface of the medium with little aerial mycelium (Fig. 17). On richer media, such as oatmeal agar or potato dextrose agar, the mycelium is

Fig. 15. Aphanomyces euteiches zoospore cluster (upper left), oospore (center), and hyphae (right). (Cour tesy D. Malvick— © APS)

Fig. 13. Oospores of Aphanomyces euteiches embedded in root tissues. (Courtesy S. Chatterton— © Her Majesty the Queen in Right of Canada, as represented by the Minister of Agriculture and Agri- Food Canada)

Fig. 16. Zoospores of Aphanomyces euteiches at the apex of a sporangium. (Cour tesy D. Malvick— © APS)

Fig. 14. Yellowing symptoms of Aphanomyces root rot progressing from the bottom of the shoot upward on infected plants.

(Courtesy S. Chatterton—© Her Majesty the Queen in Right of Canada, as represented by the Minister of Agriculture and AgriFood Canada)

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Fig. 17. Aphanomyces euteiches colony on cornmeal agar. (Courtesy D. Malvick—© APS)

tough and dense with more aerial growth. Optimal growth of mycelium occurs at 28°C, with lower and upper limiting temperatures of 4 and 32°C. Zoospores can be produced by growing A. euteiches in peptone glucose broth followed by washing the mycelial mat to remove the nutrients and then immersing it in a mineral salts solution or lake water overnight.

The pathogen can be isolated from infected plant tissue but only if isolation is attempted soon after infection and before the root cortex is completely rotted. The pathogen grows in 2–3 days from surface-disinfested roots placed on water agar or on a semiselective medium. On either medium, A. euteiches may be recognized by its sparse, arachnoid growth habit and by the characteristic appearance of the hyphae under microscopic examination (Fig. 15). The coenocytic (with no or few cross walls and multiple nuclei) hyphae are 8–12 µm in diameter.

Parts of the interior of the hyphae appear empty of content, and other parts appear filled with granular cytoplasm. Isolates of A. euteiches can be stored on cornmeal agar for 3–5 months at 4–8°C. To confirm the identity of a suspected A. euteiches isolate, a 0.5-cm 2 piece of agar containing mycelium from a 3- to 4-day-old culture can be placed in sterile tap water. Characteristic primary spore clusters and zoospores are then observed around the edges of the agar within 1–3 days. Pathogenicity can be tested by inoculating pea seedling roots with a zoospore suspension and observing symptom and oospore development in roots after 7–14 days of incubation.

A. euteiches can be isolated from infested soil by a rolledtowel assay (soil and seeds rolled in sterile paper towels or germination paper and kept moist) or by baiting from soil by planting a susceptible pea cultivar in pots containing infested soil. Quantitative real-time PCR can be used with root and soil DNA extracts to quantify pathogen biomass and oospore levels.

A. euteiches has a wide host range; however, individual isolates can be host selective. The host range in controlled inoculation assays includes pea, alfalfa, snap bean, dry bean, soybean, red clover, lentil, faba bean, cicer milkvetch (a species of Astragalus), and many different weed species. A few nonlegumes, such as spinach, oat, and redroot pigweed, can support infection and oospore production by A. euteiches. Populations of A. euteiches strains vary in host preference and virulence.

Pea-infecting isolates of A. euteiches may infect other legumes, including alfalfa and lentil. Two main pathotypes have been reported in pea on the basis of their differential reactions on a set of six genotypes, especially the MN313 pea germplasm line.

Pathotype I is predominant in Europe and was observed in the United States, and pathotype III was detected at some locations in the United States. Most isolates from bean and red clover do not infect pea. Genotypic analysis with neutral DNA markers has detected genetic distinctions among strains isolated from different hosts, fields, regions, and/or countries. Variation in aggressiveness on pea genotypes has also been reported among A. euteiches populations from France and the United States.

Disease Cycle and Epidemiology A. euteiches can infect pea plants at any stage of growth, although the most damage and yield loss occur when plants are infected at young, seedling growth stages. Infection generally occurs in the month following emergence in heavily infested fields. Aboveground symptoms may appear as early as 10 days after emergence if the inoculum level is high and the soil is wet and warm. Symptoms appear later in the season when inoculum levels are low or weather conditions are less conducive to disease development.

Infection begins when zoospores or oospores germinate on the surfaces of roots and the mycelium penetrates and grows through the host tissue. Enzymes that degrade root tissue are then released by the pathogen. Mature oospores are formed within the root 1–2 weeks after infection. The oospores are released into the soil as the roots decompose or remain inside root fragments, and they serve as inoculum during subsequent years. Oospores are found primarily in the plowed layer of soil to a depth of 60 cm, where they may persist for 10 years or longer. In no-till systems, oospore levels are highest in the top 0–20 cm of soil but can also be found deeper (20–60 cm). The pathogen can move a short distance within a field during a growing season, presumably by means of mycelial growth between adjacent roots or by movement of zoospores. A. euteiches can also move long distances with infested soil and vines or during periods of flooding and along water tracks (Fig. 18). The pathogen tends to be distributed across fields, especially in fields that are highly infested and where complete yield loss occurs. In low to moderately infested fields, disease foci may be present and yellow patches can develop (Fig. 19). Alternative hosts and susceptible weed species can promote survival and reproduction of this pathogen.

Infection occurs at all temperatures conducive to pea growth; however, symptoms develop most rapidly at 22–28°C. Wet, slowly drained, and compacted soil favors infection and disease development. Warm periods with low rainfall accentuate

Fig. 18. Aphanomyces euteiches can move with water and cause root rot across entire fields. (Courtesy S. Chatterton—© Her Majesty the Queen in Right of Canada, as represented by the Minister of Agriculture and AgriFood Canada)

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the water stress experienced by infected plants, and Aphanomyces root rot can be especially severe during years when a wet and warm spring is followed by dry, warm weather. The disease is not limited to soils with a high water-holding capacity and can be severe on sandy soils kept moist by irrigation or frequent rain events. In the Canadian prairies, disease prevalence and incidence are similar across all of the three major soil zones where peas are grown, despite inherent differences in soil water-holding capacity. In France, the disease incidence is similar in different soil types, except that soils rich in limestone are less prone to the disease.

Because pathogen inoculum levels can increase quickly when pea is grown in infested soil and decrease slowly even in the absence of a suitable host, this disease is difficult to manage. The rate of inoculum decline depends on soil type, crop rotation, and other factors, but highly infested fields can remain risky for pea production after 8 years or more of nonhost crops.

An epidemic arising from a low inoculum level and causing only slight aboveground symptoms can produce sufficient inoculum to destroy the subsequent pea crops if conditions favor the disease. An inoculum density of 100 oospores per gram of soil is sufficient to cause moderate to severe root rot (Fig. 20).

Although damage caused by A. euteiches on its own can be very severe, pea roots are seldom infected by this pathogen alone. Other soilborne organisms are generally part of the disease complex. When A. euteiches is present at low or moderate

Fig. 19. Early in the establishment of root rot caused by Aphanomyces euteiches patches (disease foci) may occur. (Courtesy S. Chatterton— © Her Majesty the Queen in Right of Canada, as represented by the Minister of Agriculture and Agri-Food Canada)

Fig. 20. Dilution curve of Aphanomyces euteiches oospores from 0 to 5,000 (right to left) causing root symptoms on pea. (Cour tesy S. Chatterton— © Her Majesty the Queen in Right of Canada, as represented by the Minister of Agriculture and Agri- Food Canada)

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inoculum levels, infection of roots by pathogens such as Fusarium or Pythium spp. or by nematodes such as Pratylenchus spp. can increase disease severity. Viral infections can also increase the severity of Aphanomyces root rot.

Management Research to manage Aphanomyces root rot has been ongoing since this disease was first described in 1924. Despite many attempts with fungicides, cultural practices, and host resistance, Aphanomyces root rot remains very difficult to manage and is destructive in many pea-production areas. Encouraging but inconsistent results have been obtained with experimental fungicides, fertilizers, or mineral salts. Ethaboxam is the only component currently registered for early season suppression of Aphanomyces root rot in some countries. Dinitro herbicides decreased disease severity in some greenhouse and field tests, but the resulting yield increases were generally insufficient to justify use for disease control. Precrops and cover crops, such as members of Brassicaceae or oat, also have been reported to suppress the disease; however, these practices are not always effective and not widely used. For example, oat planted prior to pea can potentially reduce the severity of Aphanomyces root rot when oat plants are chisel plowed in the previous fall after being grown as a summer crop. Soil compaction can enhance Aphanomyces root rot and should be minimized.

An effective way to reduce losses caused by this disease is to check fields for infestation level (root rot inoculum potential (IP)) before planting and to avoid moderately to highly infested fields. The routine use of such a program has been of economic benefit to pea growers. For example, in France, a test has been developed to evaluate the A. euteiches IP level of the soil on a 0–5 scale. Recommendations are then given to growers that are based on the IP level of the field. When the IP is low, spring pea cultivars that are susceptible to the disease should not be grown. When the IP is moderate, winter pea growing is possible because the older plant stage at the time of infection in the spring makes it less susceptible to the disease than spring pea. When the IP is high, no peas should be grown. A similar soil-testing and recommendation system has been used in the United States. Laboratories in Canada also offer soil-testing services for the presence or absence of A. euteiches based on DNA screening of soil samples, but to date they do not provide information on IP. Crop rotation with nonhost crops may reduce inoculum potential of A. euteiches, although many years away from susceptible crops may be needed to substantially reduce the IP. Numerous legume species can be hosts, and unless they have been tested, they should be avoided. For example, soybean, lupine, chickpea, fenugreek, and lotus are resistant and many cultivars of faba bean are resistant. On the other hand, lentils, clovers, and alfalfa are susceptible to the disease and contribute to increases in IP in soil, although some cultivars are less susceptible than others.

To date, there are no commercial cultivars of pea available with high levels of resistance to Aphanomyces root rot.

However, progress is being made in this regard. Pea lines partially resistant to A. euteiches were identified from germplasm screening and breeding programs conducted in the United States and France. Two of seven identified genomic regions, Ae-Ps7.6 and Ae-Ps4.5, had a major effect on resistance to A. euteiches strains of pathotypes I and III, respectively. The major quantitative trait locus (QTL), Ae-Ps7.6, was associated with resistance and root architecture traits and reduced root losses caused by infection. The research advances in genetics have provided QTL-linked markers and partially resistant germplasm, which are being used to develop pea cultivars with increased resistance to Aphanomyces root rot.

In summary, Aphanomyces root rot is a destructive disease of pea that should be managed by combining several methods.

These include avoidance of heavily infested land, prevention

of inoculum buildup through the use of long rotations with nonhost plants, the use of resistant crops or those that escape disease through the timing of planting, and the use of partially resistant pea cultivars when available.

Selected References Banniza, S., Bhadauria, V., Peluola, C. O., Armstrong- Cho, C., and Morrall, R. A. A. 2013. First report of Aphanomyces euteiches in Saskatchewan. Can. Plant Dis. Surv. 93:163-164.

Chatterton, S., Bowness, R., and Harding, M. W. 2015. First report of root rot of field pea caused by Aphanomyces euteiches in Alberta, Canada. Plant Dis. 99:288.

Desgroux, A., Baudais, V., Aubert, V., Le Roy, G., de Larambergue, H., Miteul, H., Aubert, G., Boutet, G., Duc, G., Baranger, A., Burstin, J., Manzanares-Dauleux, M., Pilet-Nayel, M.-L., and Bourion, V. 2018.

Comparative genome-wide-association mapping identifies common loci controlling root system architecture and resistance to Aphanomyces euteiches in pea. Front. Plant Sci. 8:2195.

Fritz, V. A., Allmaras, R. R., Pfleger, F. L., and Davis, D. W. 1995.

Oat residue and soil compaction influences on common root rot (Aphanomyces euteiches) of peas in a fine-textured soil. Plant Soil 171:235-244.

Gangneux, C., Cannesan, M.-A., Bressan, M., Castel, L., Moussart, A., Vicré-Gibouin, M., Driouich, A., Trinsoutrot-Gattin, I., and Laval, K. 2014. A sensitive assay for rapid detection and quantification of Aphanomyces euteiches in soil. Phytopathology 104:1138-1147.

Grau, C. R., Muehlchen, A. M., Tofte, J. R., and Smith, R. R. 1991.

Variability in virulence of Aphanomyces euteiches. Plant Dis. 75:1153-1156.

Grünwald, N. J., and Hoheisel, G.-A. 2006. Hierarchical analysis of diversity, selfing, and genetic differentiation in populations of the oomycete Aphanomyces euteiches. Phytopathology 96:1134-1141.

Heyman, F., Lindahl, B., Persson, L., Wikström, M., and Stenlid, J. 2007. Calcium concentrations of soil affect suppressiveness against Aphanomyces root rot of pea. Soil Biol. Biochem. 39:2222-2229.

Hossain, S., Bergkvist, G., Glinwood, R., Berglund, K., Mårtensson, A., Hallin, S., and Persson, P. 2015. Brassicaceae cover crops reduce Aphanomyces pea root rot without suppressing genetic potential of microbial nitrogen cycling. Plant Soil 392:227-238.

Kraft, J. M., Marcinkowska, J., and Muehlbauer, F. J. 1990. Detection of Aphanomyces euteiches in field soil from northern Idaho by a wet-sieving/baiting technique. Plant Dis. 74:716-718.

Lavaud, C., Baviere, M., Le Roy, G., Hervé, M., Moussart, A., Delourme, R., and Pilet-Nayel, M.-L. 2016. Single and multiple resistance QTL delay symptom appearance and slow down root colonization by Aphanomyces euteiches in pea near isogenic lines.

BMC Plant Biol. 16:166.

Le May, C., Onfroy, C., Moussart, A., Andrivon, D., Baranger, A., Pilet-Nayel, M.-L., and Vandemark, G. 2018. Genetic structure of Aphanomyces euteiches populations sampled from United States and France pea nurseries. Eur. J. Plant Pathol. 150:275-286.

Malvick, D. K., and Percich, J. A. 1998. Genotypic and pathogenic diversity among pea-infecting strains of Aphanomyces euteiches from the central and western United States. Phytopathology 88:915-921.

Malvick, D. K., Grünwald, N. J., and Dyer, A. T. 2009. Population structure, races, and host range of Aphanomyces euteiches from alfalfa production fields in the central USA. Eur. J. Plant Pathol.

123:171.

Malvick, D. K., Percich, J. A., Pfleger, F. L., Givens, J., and Williams, J. L. 1994. Evaluation of methods for estimating inoculum potential of Aphanomyces euteiches in soil. Plant Dis.78:361-365.

Mieuzet, L., Quillévéré, A., Pilet, M. L., and Le May, C. 2016. Development and characterization of microsatellite markers for the oomyceta Aphanomyces euteiches. Fungal Genet. Biol. 91:1-5.

Moussart, A., Even, M.-N., and Baranger, A. 2016. Effect of pea sowing date on Aphanomyces root rot development and yield losses.

Poster presented at the International Legume Society Congress.

Moussart, A., Even, M. N., Lesné, A., and Tivoli, B. 2012. Successive legumes tested in a greenhouse crop rotation experiment modify the inoculum potential of soils naturally infested by Aphanomyces euteiches. Plant Pathol. 62:545-551.

Moussart, A., Even, M.-N., Pilet-Nayel, M.-L., and Baranger, A. 2017.

Agricultural practices to prevent and reduce Aphanomyces root rot

epidemics and damages on the pea crop in France. Page 21 in: Proc.

Int. Legume Root Dis. Workshop, 7th.

Moussart, A., Even, M. N., and Tivoli, B. 2008. Reaction of genotypes from several species of grain and forage legumes to infection with a French pea isolate of the oomycete Aphanomyces euteiches. Eur. J. Plant Pathol. 122:321-333.

Moussart, A., Wicker, E., Duparque, M., and Rouxel, F. 2001. Development of an efficient screening test for pea resistance to Aphanomyces euteiches. Pages 272-273 in: 4th European Conference on Grain Legumes—Towards the Sustainable Production of Healthy Food, Feed and Novel Products. Association européenne de recherche sur les protéagineux, Paris.

Moussart, A., Wicker, E., Le Delliou, B., Abelard, J.-M., Esnault, R., Lemarchand, E., Rouault, F., Le Guennou, F., Pilet-Nayel, M.-L., Baranger, A., Rouxel, F., and Tivoli, B. 2009. Spatial distribution of Aphanomyces euteiches inoculum in a naturally infested pea field.

Eur. J. Plant Pathol. 123:153-158.

Quillévéré-Hamard, A., Le Roy, G., Moussart, A., Baranger, A., Didier, A., Pilet-Nayel, M.-L., and Le May, C. 2018. Genetic and pathogenicity diversity of Aphanomyces euteiches populations from pea-growing regions in France. Front. Plant Sci. 9:1673.

Vandemark, G., and Grünwald, N. J. 2005. Use of real-time PCR to examine the relationship between disease severity in pea and Aphanomyces euteiches DNA content in roots. Eur. J. Plant Pathol.

111:309-316.

Wicker, E., and Rouxel, F. 2001. Specific behaviour of French Aphanomyces euteiches Drechs. populations for virulence and aggressiveness on pea, related to isolates from Europe, America and New Zealand. Eur. J. Plant Pathol. 107:919-929.

Williams-Woodward, J. L., Pfleger, F. L., Fritz, V. A., and Allmaras, R. R. 1996. Green manures of oat, rape, and sweet corn for reducing common root rot in pea (Pisum sativum) caused by Aphanomyces euteiches. Plant Soil 188:43- 48.

Zitnick-Anderson, K., and Pasche, J. S. 2016. First report of Aphanomyces root rot caused by Aphanomyces euteiches on field pea in North Dakota. Plant Dis. 100:522.

Prepared by W. Pfender, D. K. Malvick, F. L. Pfleger, and C. R. Grau; revised by M.-L. Pilet-Nayel, A. Moussart, S. Chatterton, and D. K. Malvick

Fusarium Root Rot Fusarium root rot was first described as a serious disease of pea in North America and Europe more than a century ago and has become an important root disease across the pea-producing areas of North America in both dryland and irrigated areas.

Yield losses of over 50% have been reported, but they can be even higher when other pathogens are present. Fusarium root rot often occurs in conjunction with Aphanomyces, Rhizoctonia, or Pythium root rots. Several Fusarium species have been associated with the Fusarium root rot complex; however, F. solani and F. avenaceum are the most damaging in North America. F. solani is recognized as a species complex and has historically been considered the principal pathogen of pea. However, F. avenaceum has been identified as causing major root rot on pea in Montana and North Dakota in the United States and in Canada and has a wide host range that includes numerous rotational crops (e.g., lentil, wheat, barley, and canola) in the northern Great Plains of North America.

Symptoms Infection often occurs at the cotyledonary attachment area below ground near the epicotyl, hypocotyl, or the upper taproot (Fig. 21). Lesions extend upward to the soil line and downward into the root zone. Initial symptoms on primary and secondary roots consist of reddish brown to black lesions that coalesce during the season. A red discoloration of the root vascular system can be observed, especially near the area of cotyledon attachment, but generally it does not progress above the soil line, 17